Carbon and nitrogen turnover in the Arctic deep sea: in situ benthic community response to diatom and coccolithophorid phytodetritus

Carbon and nitrogen turnover in the Arctic deep sea

Carbon and nitrogen turnover in the Arctic deep sea: in situ benthic community response to diatom and coccolithophorid phytodetritusCarbon and nitrogen turnover in the Arctic deep seaUlrike Braeckman et al.

In the Arctic Ocean, increased sea surface temperature and sea ice retreat
have triggered shifts in phytoplankton communities. In Fram Strait,
coccolithophorids have been occasionally observed to replace diatoms as the
dominating taxon of spring blooms. Deep-sea benthic communities depend
strongly on such blooms, but with a change in quality and quantity of
primarily produced organic matter (OM) input, this may likely have
implications for deep-sea life. We compared the in situ responses of Arctic
deep-sea benthos to input of phytodetritus from a diatom
(Thalassiosira sp.) and a coccolithophorid (Emiliania huxleyi) species. We traced the fate of 13C- and
15N-labelled phytodetritus into respiration, assimilation by
bacteria and infauna in a 4-day and 14-day experiment. Bacteria were key
assimilators in the Thalassiosira OM degradation, whereas
Foraminifera and other infauna were at least as important as bacteria in the
Emiliania OM assimilation. After 14 days, 5 times less carbon and
3.8 times less nitrogen of the Emiliania detritus was recycled
compared to Thalassiosira detritus. This implies that the
utilization of Emiliania OM may be less efficient than for
Thalassiosira OM. Our results indicate that a shift from
diatom-dominated input to a coccolithophorid-dominated pulse could entail a
delay in OM cycling, which may affect benthopelagic coupling.

The Arctic is warming more rapidly than the global average
(IPCC, 2014) with an increase in air temperature of roughly
2 ∘C since 1900. In the Arctic Ocean, this has led to a
concomitant rise in sea surface temperature of 1.5 ∘C, most
distinctly since the 1980s (Polyakov et al.,
2013). As a result, summer sea-ice extent is presently decreasing at a rate
of more than 10 % per decade (Comiso, 2010).

Fram Strait, located in the transition zone between the northern North
Atlantic and the central Arctic Ocean, is the sole deep gateway where these
two hydrographic regimes partly converge. In the eastern Fram Strait, the West
Spitsbergen Current (WSC) transports relatively warm (2.7–8 ∘C)
Atlantic water into the Arctic Ocean, while in the western strait, the East
Greenland Current (EGC) carries colder (−1.7–0 ∘C) polar water in
the upper 150 m towards the south (Nöthig et al., 2015). During the last
decade, the mean temperature of Atlantic water entering the Arctic Ocean with
the WSC increased by more than 0.05 ∘C yr−1
(Beszczynska-Möller et al., 2012).

Phytoplankton size and community structure are directly influenced by
temperature, and smaller temperate species may become established within the
phytoplankton community in areas with increased temperatures and less sea ice
(Hilligsøe et al., 2011; Morán et al., 2010). This phenomenon also
occurred in the eastern Fram Strait, where previously, phytoplankton
communities were typically dominated by diatoms, mainly
Thalassiosira spp. (Bauerfeind et al., 2009; Lalande et al., 2011).
However, during recent warmer years, phytoplankton blooms became more mixed
with Phaeocystis pouchetti (Nöthig et al., 2015; Soltwedel et
al., 2016). Also, Emiliania huxleyi (Prymnesiophyceae)-dominated
coccolithophorid blooms have been observed between 2000 and 2005 –
especially in 2004 – which has been attributed to northward transport of the
species into Fram Strait by means of the North Atlantic Current and WSC. This
“Atlantification” with a combined change in water temperature and water
mass origin has been suggested as one possible scenario for a community shift
in phytoplankton communities in Fram Strait (Bauerfeind et al., 2009).

As the recycling of the phytoplankton bloom mainly occurs in surface waters,
on average only a small fraction of the primary produced organic carbon
(< 5 %) is exported to the deep sea (Gooday and
Turley, 1990; Schlüter et al., 2000). Earlier observations in Fram
strait have shown that the particulate organic carbon (POC) export flux at
300 m is very similar to the diffusive oxygen uptake in sediments
(Bauerfeind et al., 2009), which suggests that, once passed
through the photic zone of the surface waters, degradation in the aphotic
zone of the pelagic is negligible (Cathalot et al., 2015). The
deposition of phytodetritus from surface water primary production is of
crucial importance for the deep-sea benthos (Boetius et al., 2013; Graf, 1989) that
profoundly depends both on the quality and quantity of the settling food
(Billett et al.,
2010; Ruhl and Smith, 2004; Smith et al., 2013). It is expected that a
change in quality and quantity of primarily produced organic matter (OM)
input may have implications for deep-sea life.

A powerful approach to quantify the processing of a food source in benthic
food webs is to label the OM with stable carbon and/or nitrogen isotopes.
This technique allows the fate of the food source to be traced to different
carbon and/or nitrogen pools representative of different biological
processes and groups of organisms. This approach has been successfully
applied in a wide range of study areas, from temperate, estuarine ecosystems
to the abyssal sea floor (Woulds et al., 2009,
2016). However, most studies target assimilation by specific size classes of
organisms, e.g. macrofauna, or specific taxonomic groups such as
Foraminifera (Enge et al., 2011; Nomaki et al., 2005) or Nematoda
(Guilini et al., 2010; Ingels et al., 2010) and
only few studies consider other mineralization pathways (e.g. respiration)
(Bühring et al., 2006a; Evrard et al., 2012; Woulds et al., 2016).

Experimental food web studies in the Arctic deep sea with labelled food
sources are scarce and have focussed on uptake of DOC, bacteria or diatom
detritus by nematodes (Guilini et al., 2010; Ingels et al., 2010) or
phytoplankton and ice algae by macrofauna (Mäkelä et al., 2017). The
latter studies are ship-based experiments, an approach which can involve
decompression of the samples and is therefore prone to introducing biases,
e.g. by changing the activity of the benthic biota. These types of
studies show that infauna selects its
food sources specifically or is indifferent: macrofauna seems to have a
preference for ice algae over phytoplankton in shallower Arctic water
(McMahon et al., 2006; Sun et al., 2007) but displays a dietary plasticity in
Arctic deep-sea sediments, assimilating both ice algae and phytoplankton
efficiently (Mäkelä et al., 2017). Also, in the abyssal Pacific,
macrofauna did not show a preference for any type of food source when
phytodetritus was readily available, whereas foraminifera selected
coccolithophore nitrogen over diatom nitrogen (Jeffreys et al., 2013).

In this study, we compare the in situ mineralization pathways of
phytodetritus of a traditionally prevailing primary producer with a food
source that will possibly dominate in the near future as a consequence of
global change effects on phytoplankton communities. Such a shift may entail
significant effects on the ecosystem, since the availability and degradation
patterns of different food sources might differ (Mäkelä et al., 2017;
Ruhl and Smith, 2004; Smith et al., 2013). One mechanism would be the
impediment of food source utilization by the physical protection of the
cells. The previously dominant diatom Thalassiosira, for example, is
protected by a silica wall which can be easily dissolved by bacteria (Bidle
and Azam, 1999). In contrast, the coccolithophorid Emiliania. huxleyi consists of a substantial part of inorganic carbon (coccoliths),
which can form a physical barrier to bacterial degradation (Engel et al.,
2009) and produces DMSP-related substances that act as anti-grazing compounds
for zooplankton (Hansen et al., 1996), both characteristics that could imply
a delay in its decomposition. However, E. huxleyi also contains
comparatively high levels of n−3 polyunsaturated fatty acids, essential for
growth and reproduction of eukaryotic consumers (Pond and Harris, 1996). This
high nutritional value has been used to explain the higher survival rate of
planktonic foraminifera (Anderson et al., 1979) and egg production by
calanoid copepods (Neystgaard et al., 1997) compared to when these
organisms were fed a diatom diet. Therefore, we hypothesize that a shift in
phytoplankton communities from diatoms to coccolithophorids could have
implications for the dynamics and efficiency of OM assimilation by the
deep-sea benthos and for mineralization at the Arctic deep-sea floor.

To test this, we conducted a comparative in situ experiment at the
Arctic deep-sea floor at 2500 m water depth, where we provided
13C-
and 15N-labelled phytodetritus of either the diatom
Thalassiosira sp. or the coccolithophorid Emiliania huxleyi
to the benthic community. We tracked the degradation and processing pathways
during a short- (4 days) and long-term (14 days) experiment in bacteria,
infauna (> 250 µm), and dissolved pools of inorganic
carbon and nitrogen in the pore water and the overlying water. These
experiments allow us to evaluate the general response of Arctic deep-sea
communities to food pulses as well as specific effects of changes in
phytodetritus composition.

2.1 Study site

This study was conducted during RV Maria S. Merian expedition MSM29
in June–July 2013 at Station S2 (MSM29/423-4 and MSM29/435-1),
78.776∘ N 5.2571∘, E; 2500 m water depth) of the long-term
deep-sea observatory HAUSGARTEN in Fram Strait. The deep waters of the area
are characterized by a constant temperature of −0.7 ∘C and
salinity of 35 (Beszczynska-Möller et al., 2012). The annual spring
phytoplankton bloom in 2013 occurred in June (NASA Ocean Color,
https://oceancolor.gsfc.nasa.gov/, last access: 5 March 2018) but an
earlier bloom in April had arrived at the sea floor with
9 mg POC m−2 d−1 1 month before the start of the experiment
(Salter et al. unpublished results). This implies that the benthic
communities had been able to graze on some fresh OM prior to our experiment,
but not yet on the main annual food pulse. The observed response rates are
therefore assumed to be representative rates of the processing of freshly
deposited phytodetritus.

2.2 Culturing of algae

The diatom Thalassiosira sp. and the coccolithophorid
Emiliania huxleyi were cultured at 15 ∘C (continuous light)
in artificial seawater amended with f∕2 medium (Guillard, 1975),
13C-bicarbonate, and 15N-nitrate (99 atom %
13C-enriched NaHCO3 and Na15NO3; Cambridge
Isotope Laboratories). The algal material was harvested by centrifugation
(3500–5500× g, 10–20 min). The harvest was frozen at
−26 ∘C. Thalassiosira contained
33 atom % 13C and 35 atom % 15N, while
Emiliania contained 31 atom % 13C and 43 atom %
15N (measured via elemental analyser – isotope ratio mass
spectrometry – EA–IRMS; see below). Their corresponding molar C : N ratio
was 13.3 (Thalassiosira) and 12.5 (Emiliania). Total carbon
(TC) and organic carbon content (TOC) of the cultures were measured on
subsamples of parallel cultures that were unmodified (TC) or acidified (TOC)
prior to measurement using EA–IRMS. The corresponding TOC
content of the algae was 78 % of TC (Emiliania) and 95 % of
TC (Thalassiosira). Before use, thawed algae were washed 3 times in
0.2 µm filtered seawater to remove labelled dissolved inorganic
carbon (DIC) and dissolved inorganic nitrogen (DIN) that may have stuck to
the exterior of the organisms. These washing steps most likely also entailed
a loss of DOM including dissolved organic nitrogen (DON). DOC and TDN (i.e.
DIN from remains of culturing medium and DON from cell leakage upon thawing)
were measured in the supernatant from the three washes with a TOC-VCPH Shimadzu
instrument (Stubbins and Dittmar, 2012) with a precision better than 3 %.
In terms of carbon, on average 10.4 % of the harvested algae was lost as
DOC, whereas in terms of nitrogen, on average 53.8 % of the harvested
algae was lost as TDN. The amount of phytodetritus reported below is
corrected for this DOM loss. Although the algae were not cultured in axenic
conditions, fatty acid (FA) analysis of the algal material did not show
the presence or labelling of bacteria-specific iso- and anteiso-branched
phospholipid-derived FAs (PLFAs i14:0, i and ai15:0, i16:0), which
suggests that the bacterial contamination of the algae cultures was
negligible.

2.3 Experimental set-up

In June 2013, two landers equipped with three benthic chambers
(20 cm × 20 cm) were deployed at the deep-sea floor. One lander
deployment lasted for 4 days, and the other one lasted for 14 days to
investigate the temporal evolution of the OM processing. After arrival of the
lander systems at the sea floor, a motorized drive lowered the chambers
approx. 10 cm into the sediment to enclose 400 cm2 of sediment
together with approx. 20 cm of overlying water. Labelled algae suspensions
were added to two of the chambers of each experiment by means of an automated
dispenser (Witte et al., 2003b). A total of
69 mmol Corg m−2 (2.9 mmol N m−2) of
Thalassiosira sp. was added to one of the chambers, while 37 mmol
Corg m−2 (2 mmol N m−2) of Emiliania huxleyi was added to a second chamber. The third chamber was left unamended
(control). These additions correspond to 0.85 g Corg m−2
(0.90 g C m−2; 0.04 g N m−2) for Thalassiosira and
0.45 g Corg m−2 (0.59 g C m−2; 0.03 g N m−2)
for Emiliania. The treatments are further referred to as
“Thalassiosira”, “Emiliania” and “control”. The
amount of added carbon was aimed to simulate the annual carbon deposition in
the area. However, logistical issues with cultivation of Emiliania
unfortunately compromised the addition of a C dose comparable to
Thalassiosira. The additions correspond to 29 %–43 %
(Thalassiosira) and 15 %–23 % (Emiliania) of the
average total annual carbon deposition in the area (around
2–3 g Corg m−2) (Bauerfeind et al., 2009; Soltwedel et
al., 2016) and represent 4.5 (Emiliania) to 8
(Thalassiosira) times the amount of POC that had already arrived
during the first sedimentation peak in late May 2013.

To verify the release of algae at the sea floor, small Teflon-coated
laboratory stirring bars had been added to each algae suspension, which were
recovered together with the incubated sediments after deployment. During
the course of the incubation, the overlying water was continuously stirred
to mimic the hydrodynamic conditions. An O2 optode (type 4330,
Aanderaa, Bergen, Norway) attached to the upper lid of the chamber monitored
oxygen concentration and temperature in the overlying water. An automated
syringe sampler took 50 mL subsamples of the overlying water at seven
regularly spaced time points by means of glass syringes. At the end of the
incubation, the incubated sediments were enclosed from below with a motorized
lid before the chambers were retracted from the sea floor. Upon acoustic
release from the ship, the lander including the three chambers with the
enclosed sediment and the overlying water was recovered. On board, the height
of the overlying water body was measured with a ruler at ∼8 positions
within each chamber. This allowed the estimation of the volume of overlying
water that is necessary to calculate the fluxes at the sediment–water interface.

Due to logistic and technical constraints associated with in situ
experiments in the deep sea, our experiments are unreplicated. We have
chosen to invest in observing temporal dynamics (short- vs. long-term
incubations), which would allow us to unravel time-dependent aspects of the
mineralization pathways (carbon respiration and nitrogen mineralisation for
different food web compartments) and link these with degradation processes.
This also gave us the chance to look into the effect of different food
sources, which is highly relevant in light of the expected changes in
primary-producer communities. We acknowledge that spatial variability may
have contributed to differences observed in the algae and time treatments.
However, we assumed that the reliability of our observations on the effect
of the algae species were supported when differences between algae
treatments observed in the short incubation were confirmed by the long
incubation. A similar approach was applied in earlier comparable deep-sea
(lander) experiments, which provided valuable new insights
(Guilini et al., 2010; Woulds et
al., 2007).

2.4 Sampling

Upon lander retrieval, the syringe samples of the overlying water and the
sediments were immediately processed. Subsamples of the water samples in the
syringes were transferred to 12 mL exetainers and fixed with
100 µL HgCl2 for later analysis of 13C-DIC and 15N-DIN. Another 1.5 mL subsample
was transferred to Zinsser vials and fixed with 25 µL HgCl2
for total DIC analysis. The samples were stored at 4 ∘C until
analysis. The remaining volume was stored frozen for nutrient analysis
(without filtration; however, suspended matter in these overlying waters is
negligible).

The sediments were subsampled at a vertical resolution of 0–1, 1–2, 2–3
and 3–5 cm and each horizon was carefully mixed prior to subsampling. For
pore water analysis of 13C-DIC and 15N-DIN, 5 mL of the
sediment from each horizon was added to 12 mL exetainers pre-filled with
He-purged water. These samples were fixed with 100 µL HgCl2.
For total DIC and DIN analysis, pore water was extracted by centrifugation
and stored as described for the overlying water samples.

Due to tilting of the 4-day incubation lander by the in situ
microprofiler, the Emiliania chamber contained less sediment and
only the 0–1 cm sediment horizon could be sampled. As will be discussed
further, this missing representation of the subsurface sediments in the
4-day Emiliania chamber results in an underestimation of the
labelled phytodetritus with < 10 % recovered as total processed
carbon and uncharacterized OM.

2.5 13C-DIC analyses

A 1.5 mL water subsample was transferred to a 12 mL exetainer and degassed
with helium. To this 150 µL of 20 % H3PO4 was
added and the samples were left overnight so that the DIC in solution would
fill the headspace in the form of CO2. This headspace was then
measured 8 times and the stable isotope ratio
(13C∕12C) was determined via a GasBench II (Thermo
Electron, Bremen Germany) coupled to an IRMS (Thermo Quest Delta Plus, Thermo
Electron, Bremen Germany). CO2 was used as a reference gas and
bicarbonate standards with concentrations similar to that of the samples were
also added as reference. The standard deviation of the measurements was
< 0.001 atom %. To determine carbon respiration, the change in
the stable isotope ratio (excess atom % 13C) from the initial
time point was multiplied by the total DIC concentration measured by flow
injection analysis with conductivity detection (Hall and Aller, 1992)
with a precision better than 2 %.

2.6 15N-DIN analyses

Subsamples (overlying water is 4 mL, pore water is 2 mL diluted with 2 mL
milli-Q) were transferred to exetainers and degassed with helium.
15NH4+ was oxidized with hypobromite to N2
(Preisler et al., 2007; Warembourg, 1993). A second set of 4.5 mL subsamples
was also transferred to exetainers and 15NOx-
(15NO2-+15NO3-) concentrations were
determined after conversion to N2. 15NO3- was
reduced to 15NO2- using spongy cadmium, followed by
15NO2- conversion to N2 using sulfamic acid
(Füssel et al., 2012). The stable isotope ratios of 28N2,
29N2 and 30N2 were analysed by a GC–IRMS (VG
Optima, Micromass, Manchester, UK). The standard deviation of the
measurements was < 0.001 atom %. Concentrations and rates
of 29N2 and 30N2 production were calculated
from the excess relative to air, as explained in detail in Holtappels et
al. (2011), and the efficiency of 15NOx- or
15NH4+ conversion to N2 was verified using
known concentrations of 15NH4+ or 15NO3-.

2.7 General sediment analyses

Sediment median grain size was measured by laser diffraction using a Malvern
Mastersizer 2000G, hydro version 5.40. Pigments were extracted with 90 %
acetone and measured with a TURNER fluorimeter (Holm-Hansen et al., 1965;
Yentsch and Menzel, 1963). TOC and TN were measured with an elemental analyser after acidification of the sediments to remove inorganic carbon in
aliquots of approx. ∼ 15 mg (dry weight). Another sediment subsample
was taken for stable carbon and nitrogen isotope analysis (see below).

Microbial (bacterial and archaeal) cell numbers were determined using AODC. Next 1
or 2 mL of sediment were fixed with sterile filtered formalin/seawater at a
final concentration of 2 % and stored at 4 ∘C. Samples were
processed as previously described (Hoffmann et al., 2017), and two replicate
filters were counted for each sample using an epifluorescence microscope
(Axiophot, Zeiss). Extracellular enzymatic turnover rates in the sediment
were determined on board using the fluorogenic substrate
fluorescein diacetate (FDA) as an indicator of the potential hydrolytic
activity of bacteria (Köster et al., 1991) but only from the short-term
(4 days) experiment.

2.9 DNA extraction, PCR, amplicon sequencing, and sequence processing

DNA was extracted from 0.5 g sediment using the MoBio PowerSoil Kit (MO BIO
Laboratories, Inc.) following manufacturer's instructions and eluted in a
final volume of 60 µl TE-buffer (10 mM Tris-Cl, pH 8.0, 1 mM
EDTA), instead of solution S5 provided in the kit. DNA was quantified using a
microplate spectrometer (InfiniteR 200 PRO NanoQuant, TECAN Ltd,
Switzerland). Amplicon libraries of the bacterial V4–V6 region of the 16S
rRNA gene were generated according to the protocol recommended by Illumina
(16S Metagenomic Sequencing Library Preparation, Part #15044223, Rev. B),
using the primers S-D-Bact-0564-a-S-15 and S-*Univ-1100-a-A-15 primer
(Klindworth et al., 2013). Sequencing was performed on an Illumina MiSeq 161
platform in 2×300 cycles paired end runs. Raw paired-end sequences
have been submitted to ENA under INSDC accession number PRJEB25160 (under
embargo, will be released upon acceptance using the data brokerage service of
the German Federation for Biological Data (GFBio, Diepenbroek et al., 2014).

Sequence processing included the following steps: primer sequences were
removed using cutadapt (v. 1.8.1, Martin, 2011). Forward and reverse
reads were merged using pear (v. 0.9.5, Zhang et al., 2013); all
sequences were trimmed and quality filtered using trimmomatic
(v. 0.32, Bolger et al., 2014). Reads were then clustered into OTUs by
applying a local clustering threshold of days equal to 1 and the fastidious
option in swarm (v. 2.1.1, Mahé et al., 2015). The SINA aligner
(v.1.2.10, Pruesse et al., 2012) was used to align and classify the seed
sequence of each OTU with the SILVA SSU database release 123 (Quast et al.,
2012). We removed all OTUs that were classified as chloroplasts,
mitochondria, archaea, or those that could not be classified at domain level
from further analysis. Absolute singletons, i.e. sequences occurring only
once in the entire data set, were removed from further analyses. This resulted
in a final number of 3635 bacterial OTU.

2.10 Sediment lipid extraction and FA analyses

The bacterial incorporation of added phytodetritus was estimated through the
isotope enrichment of bacterial-specific PLFAs (Boschker and Middelburg,
2002). Two main biomarkers were chosen for the analysis: iC15:0 and
ai15:0 because of their specificity for bacteria and presence in all
samples. Lipid extraction was performed using a modified method from Bligh
and Dyer (Bligh and Dyer, 1959) according to Sturt et al. (2004). In short,
lipids were extracted using a mixture of methanol, dichloromethane and
phosphate buffer to pH 7.4 or trichloroacetic acid (2:1:0.8v∕v). From
this total lipid extract, an aliquot (1∕2) was saponified using 6 % KOH
in methanol, after which neutral lipids were released with hexane and
subsequently removed (Elvert et al., 2003). The remaining methanolic water
phase was acidified to pH 1 and free FAs were extracted with hexane. FAMEs
were identified via GC–MS (Thermo Quest Trace GC with Trace MS) and
concentrations determined by gas chromatography (GC)–flame ionization
detection (Thermo Finnigan Trace GC) relative to the internal standard (IS)
2Me-octadecanoic acid added prior to extraction. Corresponding stable carbon
isotope compositions of FAMEs were determined by GC–IRMS (Thermo Scientific
V Delta Plus with Trace GC ultra, connected via GC Isolink and ConFlo IV
interfaces) using CO2 as a reference, and cross-checked against the
known δ13C value of the IS FA. δ13C values have
been corrected for the methyl group added during derivatization.
δ13C values have an analytical error of 1 ‰ based on
duplicate injection of selected samples.

2.11 Fauna analysis

Following staining with rose Bengal, sediments were sieved on a
250 µm mesh to retrieve macrofauna and the larger fraction of the
meiofauna. We did not consider the meiofauna fraction
< 250 µm, since earlier studies in the research area have
proven that nematodes constitute the bulk of this fraction and their share in
13C assimilation is
negligible (Guilini et al., 2010; Ingels et al., 2010). Organisms were sorted
at a higher taxon level (Nematoda, Foraminifera, Polychaeta, Bivalvia,
Amphipoda, Tanaidacea, Porifera) and in most cases pooled over several depth
layers to reach sufficient biomass. After drying at 60 ∘C of each
taxon fraction, samples were prepared for stable carbon and nitrogen isotope
analysis via EA–IRMS (see below). Faunal biomass was determined via C
content values from the IRMS, combined with faunal abundance.

2.12 EA–IRMS analysis

Oven-dried sediment and fauna samples were decalcified overnight with the
fumes of 37 % HCl in a desiccator. Freeze-dried algae samples were used
without decalcification. Prepared samples were packed into tin cups and
analysed by a Thermo Flash EA 1112 elemental analyser coupled to an isotopic
ratio mass spectrometer (Thermo Delta Plus XP, Thermo Fisher Scientific,
Waltham, MA, USA). Caffeine was used as a standard for isotope correction and
C : N quantification of bulk carbon and nitrogen assimilation. Precisions
of the caffeine measurements were C =1.07±4.57×10-5 atom % and N =0.37±6.22×10-5 atom %
(n= 23).

2.13 Calculations

Rates of total oxygen uptake (TOU), 13C-DIC and
15N–NOx- accumulation in the overlying water
were calculated from the slope of linear regressions of concentration as a
function of time. Only significant (p< 0.05) and linear
accumulation or consumption (checked by linear regression) of the mentioned
species was considered.

15N-NH4+ concentrations in the water column did not follow a
linear increase or decrease over time, but rather an initial increase
followed by a decrease. Therefore, the minimum turnover of
15N-NH4+ was calculated as the sum of the initial
accumulation and the following consumption in the water column.

The carbon accumulation in the DIC pool in the pore water, bulk sediment and
fauna was calculated as the product of the excess atom % 13C
and the carbon content of the sample, divided by the atom % 13C
of the labelled algae:

13C label incorporation into bacterial biomass was based on the
bacterial PLFAs that were present in all chambers and depth horizons
(i15:0, ai15:0) (Boschker and Middelburg, 2002). For each bacterial PLFA,
13C label incorporation was calculated as follows:

(2)IPLFA=EPLFA×PLFAcarbonconcentration,

where excess 13C (E) is given by the difference in fraction
13C in the sample (Fsample) and the background
(Fbackground).

(3)E=Fsample-Fbackground,

where

(4)F=13C13C+12C=RR+1,

and

(5)R=δ13C1000+1×RVPDB.

Subsequently, incorporation into bacterial biomass was calculated based on
Middelburg et al. (2000) as follows:

(6)I=sumIPLFAa×b,

where a is the average PLFA concentration in bacteria (0.073 g
PLFA C g−1 of C biomass in oxidized sediments (Brinch-Iversen and King,
1990) and b is the fraction of the bacterial PLFA considered here that are
encountered in sediments of HAUSGARTEN (0.14; calculated from the fraction of
i15:0 and ai15:0 in the control sediments of Guilini et al. (2010) and
those in the present study; data not shown). The total amount of algal C
(12C+13C) recovered from bacteria, fauna, and DIC
pools was calculated as the quotient of the total accumulation I and the
fractional abundance of C in the algae (0.31–0.33).

Figure 1Concentration of excess bulk 13C-POC (a) and
15N-PN (b) in the sediment and 13C-DIC
(c), 15N-NH4+(d) and
15N-NOx-(e) in the pore water of each
experiment. Only the first sediment centimetre could be sampled in the
Emiliania 4-day experiment.

3.2 Algae addition

The addition of 69 mmol Corg m−2Thalassiosira
detritus and 37 mmol Corg m−2Emiliania detritus
reflects an addition of 1.0 % (Thalassiosira) and 0.5 %
(Emiliania) fresh organic carbon to the background TOC pool in the
upper centimetre of the sediment. For nitrogen, this corresponds to a
0.4 % (Thalassiosira) and 0.3 % (Emiliania)
addition to the background pool.

Table 1Carbon and nitrogen stock inventories for the four experiments. Data
are shown as concentrations (mmol m−2) and as a percentage of the
amount of carbon or nitrogen added as algal detritus.

* Only the first
sediment cm could be sampled in the Emiliania 4-day experiment.

Chlorophyll a (Chl a) concentrations in the control sediments were low
(1.4 µg mL−1 in the 0–1 cm of the 4-day experiment and
0.6 µg mL−1 in the 14-day experiment). The addition of the
fresh OM was reflected in an excess in Chl a in the surface sediments
(Fig. S1 in the Supplement). This excess was especially pronounced in the
Thalassiosira experiments where the addition of phytodetritus more
than doubled the Chl a content in the uppermost centimetre compared to the
control. This increase is clearly associated with the algae addition as it
largely exceeds the natural variability among replicates of surface sediments
at the same station, which is of the order 15 % (Schewe, 2018).

3.3 Sediment POC and PN pools

After 4 days, labelled Emiliania and Thalassiosira biomass
had accumulated in the organic carbon and nitrogen pools of the upper 2 cm of the sediment (Fig. 1a, b). Accumulation of label continued in
the 14-day experiment (Fig. 1a, b). Over the time course, the subsurface
sediment layers (1–3 cm) became increasingly enriched with carbon and
nitrogen. Integrated over depth, the algae-derived matter that accumulated in
the sediments represented 30 %–60 % of the originally added
Thalassiosira Corg and 8 %–52 % of the
originally added Emiliania Corg (Table 1). In terms of
nitrogen, the accumulation represented 36 %–137 % of the originally
added Thalassiosira N and 14 %–65 % of the originally added
Emiliania N.

3.4 Pore-water-labelled DIC and DIN pools

Respiration of 13C-labelled algae released 13C-DIC to
pore waters in all four experiments, with higher amounts found in the
longer-lasting experiments (Fig. 1c). The highest pore water
13C-enrichment was found in the 14-day Thalassiosira
experiment concurrent with a strong vertical gradient in 13C-DIC.
In the 14-day Emiliania experiment, however, a considerable share
of the excess 13C-DIC was found in the deepest horizon (Fig. 1c).
Integrated over the sampled sediment depth, the algae-derived DIC made up
∼0.1 % of the added organic carbon in both 4-day experiments and
increased 10-fold to ∼1 % of the added organic carbon in both
14-day experiments (Table 1).

Ammonium release from the algae to the pore waters, followed a similar
pattern as 13C-DIC
(Fig. 1d). This release added considerably to the background ammonium pore
water pool. At the end of the 4-day and 14-day Thalassiosira
experiments, 15N-NH4+ concentrations represented
14 %–47 % of the total ammonium found in the 0–1 cm sediment layer
(compare with Fig. S2). At the end of the Emiliania experiments,
15N-NH4+ in the pore water of the 0–1 cm sediment
layer made up 4 %–18 % of the total ammonium concentration (compare
with Fig. S2).

Figure 2Accumulation of 13C-DIC (a), 15N-NH4+(b) and 15N-NOx-(c) in the
water column over time in the experiments. Only significant linear
regressions are shown in full grey lines. The dashed grey line in the
15N-NH4+ data of the Thalassiosira 4-day
experiment represents a non-significant regression based on 3 points (p=0.2).

At concentrations that were approx. 1 order of magnitude lower compared to
15N-NH4+, 15N-NOx- also
accumulated in the pore waters of the experiments. In the surface (0–1 cm)
sediment layer, these 15N-NOx- concentrations
made up < 1% of the total NOx- pool.
15N-NOx- concentrations decreased with depth in
all experiments (Fig. 1e).

In total, 0.2 %–0.6 % of the added algal nitrogen was recovered as
DIN in the pore water of the 4-day Thalassiosira and
Emiliania experiments. Algal-derived DIN continued to accumulate in
the pore water to 1 and 4 % at the end of the 14-day Emiliania
and Thalassiosira experiments, respectively (Table 1).

3.5 Labelled DIC dynamics in the overlying water

Dynamics of dissolved inorganic carbon and nitrogen species in the overlying
water reflect the release from the sediment and are presented as a measure
of the respiration and mineralization of the organic material by the
sediment community. They may, however, to some extent also represent
processes that take place within the overlying water itself.

In the first 3 days of both Emiliania incubations, 13C-DIC
concentrations quickly accumulated in the overlying water, after which the
increase levelled off (Fig. 2a). The increase in 13C-DIC in the
Thalassiosira chambers was higher and steady. The linear increases
over time correspond to 13C-DIC fluxes in Thalassiosira
chambers of 54 (4 days) and
124 µmol m−2 d−1 (14 days) and in Emiliania
chambers of 103 µmol m−2 d−1 (4 days) and
18 µmol m−2 d−1 (14 days; this linear increase was only
significant at p= 0.08). Converting it into algae-derived organic carbon
(divide the 13C-DIC fluxes by fractional abundance of C in the
algae, i.e. by 0.33 for Thalassiosira and by 0.31 for
Emiliania) results in a mineralization of 1 %–8 % of
the added Thalassiosira-Corg and 2 %–4 % of the
added Emiliania-Corg (Table 1) over the time course of
the respective deployments.

3.6 Total oxygen uptake

TOU in the control chambers as calculated from oxygen recordings of the
optodes over time ranged between 0.4
(14 days) and 1.03 mmol O2 m−2 d−1 (4 days). The
latter value probably overestimates TOU owing to the presence of a shrimp
in this particular benthic chamber. Due to the large range of oxygen uptake
rates in the control chambers and also strong variabilities in the
treatments, the effect of the addition of fresh phytodetritus on TOU is not
easily recognized: TOU in the Thalassiosira chambers ranged between
0.63 (4 days) and 1.62 mmol O2 m−2 d−1 (14 days),
whereas TOU in the Emiliania chambers ranged between 0.14 (4 days) and
0.46 mmol O2 m−2 d−1 (14 days). Not including the
exceptionally low TOU value in the Emiliania 4-day experiment,
these fluxes are within the natural variability of the TOU measured for this
area, which is in the range of 0.7–1.5 mmol m−2 d−1
(Wenzhöfer et al., unpublished data).

3.7 Labelled DIN dynamics in the water column

15N-NH4+ concentrations in the overlying water of the
Thalassiosira experiments initially increased (first 2.5 days: 7.7
and 4.3 µmol m−2 d−115N-NH4+ in
the 4 and 14-day experiments, respectively), before they started to decrease
at relatively high rates (15.9 and 1.7 µmol m−2 d−115N-NH4+ in the 4 and 14-day experiments,
respectively). The 15N-NH4+ concentrations in the
overlying water were 1 order of magnitude higher than the
15N-NOx- concentrations (Fig. 2b).
15N-NOx- concentrations steadily increased over
time in both Thalassiosira experiments (0.56 and
0.35 µmol m−2 d−115N-NOx- in
the 4 and 14-day experiments, respectively), which suggests continuous
nitrification of ammonium released from Thalassiosira phytodetritus.

Different temporal patterns were observed in the Emiliania
experiments. 15N-NH4+ concentrations in the overlying
water of the Emiliania 14-day experiment steadily decreased at a
rate of 0.9 µmol m−2 d−1, while
15N-NOx- concentrations increased at a rate of
0.38 µmol m−2 d−1.

After being converted into algae-derived nitrogen (division by fractional abundance of
0.35 for Thalassiosira and by 0.43 for Emiliania), the sum
of the NH4+ and NOx- fluxes results in a
mineralization of 4 %–11 % of the added Thalassiosira-N and
0 %–2 % of the added Emiliania-N (Table 1).

Figure 3Total assimilation of algal carbon (mmol 13C m−2) in
bacterial biomass in the sediment of the 4 and 14-day
Emiliania and Thalassiosira experiments. For the 4-day
Emiliania experiment, only the first sediment centimetre was available for
analysis.

3.8 Biotic response

3.8.1 Bacterial numbers, activity and microbial community structure

Bacterial numbers determined by means of AODC direct counts did not
increase with the addition of fresh OM (Fig. S4), but exoenzymatic
activities suggest that bacterial esterase activity in the upper 2 cm of sediment increased by 19 %–36 % in the 4-day
experiment (Fig. S5) compared to control bacterial enzymatic activity
(1–1.55 nmol mL−1 h−1 in the upper 2 cm). This excess
slightly exceeds the natural variability among replicates, which is
15 %–17 % for surface sediments at the same station (Schewe, 2018).
Bacterial enzymatic activity was not measured in the 14-day experiment.
Proteobacteria, in particular Gammaproteobacteria, dominated
bacterial communities in all samples (about 35 % of total community), but
Gammaproteobacteria had a strikingly higher proportional abundance
(∼60 %) in the Thalassiosira 14-day experiment (Fig. S6,
S7), resulting in lower diversity estimates based on the inverse Simpson
index (Table S1 in the Supplement). More specifically, Colwelliaceae and
Oceanospirillaceae were conspicuous families in sediments from this
treatment, both accounting for about 15 % of the total microbial
community, in comparison to < 2 % in all other samples. The
number of OTUs in the different samples ranged between 700 and 1000, with
highest numbers in the Emiliania 14-day treatment (Table S1).

Figure 4Algal detritus assimilation by the different infauna taxa in terms
of carbon (a) and nitrogen (b). Only the first sediment
centimetre could be sampled in the Emiliania 4-day experiment.

3.8.2 Accumulation of algal carbon in bacterial biomass

Assimilation of the added fresh organic carbon into bacterial biomass
determined via 13C uptake into bacteria-specific branched FAs
(i15:0 and ai15:0) was highest in the surface sediments and increased
over time (Fig. 3). With time, the subsurface bacteria also incorporated the
13C-labelled carbon derived from the phytodetritus. When integrated
for the full sediment column investigated, bacterial carbon assimilation
ranged from 0.3 % (Emiliania) to 1.2 %
(Thalassiosira) of the added organic carbon in the 4-day experiments
and increased to 1.6 % (Emiliania) and 5.7 %
(Thalassiosira) of the added organic carbon in the 14-day
experiments (Table 1).

3.8.3 Accumulation of algal carbon and nitrogen in infaunal biomass

Carbon assimilation by infauna > 250 µm was similar in
both 4-day experiments (0.03 mmol 13Corg m−2)
and increased over time, with the highest assimilation in the
Thalassiosira 14-day experiment (0.13 mmol
13Corg m−2) (Fig. 4). The nitrogen assimilation
also increased over time, but the patterns were different, with higher
assimilation in the Thalassiosira 4-day experiment than in the
Emiliania 4-day experiment and the strongest assimilation in the
Emiliania 14-day experiment (0.023 mmol 15N m−2).

Figure 6Processed Corg(a) and N (b) in the
different compartments. Note that bacterial assimilation was not quantified
in terms of N. Only the first sediment centimetre could be sampled in the
Emiliania 4-day experiment.

Foraminifera clearly dominated the carbon and nitrogen assimilation, followed
by polychaetes. Foraminifera also displayed the highest carbon- and
nitrogen-specific assimilation of algal detritus, at least 1
(carbon-specific) to 2 (nitrogen-specific) orders of magnitude lower than
for polychaetes (Fig. 5). The specific uptake increased with time in these
two taxa but was most pronounced in Foraminifera. Foraminifera displayed a
higher carbon- and nitrogen-specific uptake of Emiliania
phytodetritus than of Thalassiosira. Bivalvia and Nematoda, the
other two relatively abundant groups, hardly incorporated any
phytodetritus-derived carbon or nitrogen into their tissue. In total, the
infauna incorporated 0.6 %–0.8 % of the added carbon in the 14-day
experiments but up to 1.4 % and 2.9 % of the added nitrogen in the
Thalassiosira and Emiliania 14-day experiment,
respectively (Table 1).

3.9 C and N budgets

3.9.1 Carbon

In the 4-day Thalassiosira experiment, 2 % of the added algal
organic carbon was processed (Table 1). A little less than half of this
(43 %) was respired, half (50 %) was assimilated by bacteria, and
7 % had been assimilated by infauna (bacterial assimilation to infauna
assimilation ratio of 7). After 14 days, 15.5 % of the added algal organic
carbon had been processed, of which now two-thirds (59 %) had been
respired. Consequently the share of the added material that was assimilated
by bacteria (37 %) and infauna (4 %) had decreased by this time
(bacterial assimilation to infauna assimilation ratio of 9) (Table 1; Fig. 6).

Different patterns were observed in the Emiliania experiments. In
the 4-day Emiliania experiment, 4 % of the added algal organic
carbon was processed and distributed over the different pools. Most of the
processed carbon had been respired to DIC (86 %), 7 % had ended up in
infauna, and the same amount had been assimilated by bacteria (bacterial
assimilation to infauna assimilation ratio of 1) After 14 days, still only
5 % of the added algal organic carbon had been processed but the
distribution was different. A larger fraction of the processed organic carbon
was present in bacterial (29 %) and infaunal (14 %) biomass
(bacterial assimilation to infauna assimilation ratio of 2) and the portion
respired amounted to 56 %.

3.9.2 Nitrogen

In the 4-day Thalassiosira experiment, 11 % of the added algal
nitrogen was processed (Table 1). However, as biotic nitrogen assimilation
only considers infauna and not nitrogen use by bacteria, the nitrogen budget
presented should be regarded as conservative. Most of the processed nitrogen
was mineralized (88 %) and recovered as ammonium and
NOx- in the water column or in the pore water DIN pool
(5 %) and only 7% in infauna. After 14 days, 16 % of the added
algal nitrogen had been processed, of which still only 9 % was traced
back into infauna and the rest was dissolved in the overlying water pool and
pore water pool (91 %) (Table 1; Fig. 6).

In the 4-day Emiliania experiment, only 0.6 % of the added algal nitrogen was
processed. In contrast to organic carbon, a much larger share of organic
nitrogen provided was assimilated. Three-quarters were found back in infauna
(73 %) and the rest (27 %) in the pore water DIN pool, whereas no
significant DIN release to the overlying water column was observed. After 14 days, 6 % of the added algal nitrogen had been processed, and still the
fauna share was high (49 %) and the rest was mainly found back as DIN in
the overlying water column and pore water pool (51 %).

In all experiments, still a considerable amount (8 %–60 % of the
added algal organic carbon; 14 %–82 % of the added algal N) of
uncharacterized OM (bulk sedimentary 13C-POC and 15N-PON)
was left in the sediment (Table 1). This pattern was most clear in the
Thalassiosira experiments. A large fraction could not be recovered:
in particular in the 4-day experiments, 68 %–88 % of the added
algal organic carbon and 7 %–86 % of the added algal N was missing;
hence carbon and nitrogen budgets cannot be closed. An exception to this
pattern is the nitrogen budget of Thalassiosira 14 days, in which
the amount of uncharacterized organic nitrogen even exceeds the total amount
of added algal nitrogen (Table 1).

We hypothesized that a potential climate-change-related shift in
phytoplankton communities from diatoms to coccolithophorids would have
implications for bathyal benthic OM assimilation and affect the
mineralization patterns at the deep-sea floor. Our results indeed show shifts
in the importance of infaunal vs. bacterial assimilation of algal organic
carbon with a lower bacterial∕infaunal assimilation ratio in
Emiliania treatments compared to Thalassiosira. In
addition, both the cycling pathways of organic carbon and nitrogen point at a
less efficient mineralization of Emiliania detritus compared to
Thalassiosira detritus: after 14 days, 5 times less carbon and
3.8 times less nitrogen of the Emiliania detritus was recycled. This
indicates that the cycling of Thalassiosira detritus was faster
compared to Emiliania.

The added Thalassiosira and Emiliania doses differed by
35 % in terms of total carbon (46 % in terms of organic carbon)
because of methodological complications. The above-mentioned difference in OM
recycling between the phytoplankton species therefore could also be partly
driven by food quantity. Experimental studies on the effect of resource
quantity on benthic mineralization pathways indeed point at a 2–10-fold
increase in bacterial carbon assimilation and 6–36 times higher carbon
respiration with a 10-fold increase in OM dose (Bühring et al., 2006b;
Gontikaki et al., 2013; Mayor et al., 2012; van Nugteren et al., 2009).
However, the difference in added OM between the experimental treatments of
these earlier studies was 1 order of magnitude, or more, which is at least
5 times larger than the difference in dose between the
Thalassiosira and Emiliania treatments in this study.
Modelling also suggested that, in the case of food-quantity-driven alterations in
OM degradation patterns, it is rather a high POC input to the abyss that
results in a stronger role for fauna-mediated carbon cycling (Dunlop et al.,
2016), whereas we found a smaller role for fauna in the treatment with the
highest POC input (Thalassiosira). Based on both the above-mentioned
experimental and modelling studies, we do not expect large effects from the
relatively small differences in the quantity of the added
Thalassiosira and Emiliania.

4.1 Carbon respiration

The relatively low amount of algal carbon mineralized in this study (with a
maximum of 15.5 % of Thalassiosira biomass after 14 days) and
the relatively slow response to OM addition is similar to other deep-sea
studies where low temperatures and limited biomass slow down the recycling of
OM (Witte et al., 2003a; Woulds et al., 2009). A delayed response with low
activities for a few days was expected as it was also observed in other
studies in cold deep-sea ecosystems (e.g. Andersson et al., 2008 in the
Arabian Sea) and resulted in the design of the experiment with a shorter and
a longer incubation. In case the previously thawed cells continued leaking
after addition, the relatively slow response may also be explained in part
by the reduced availability of labelled DOM dispersed in the overlying water
as opposed to POC at the sediment surface. The share of organic matter
provided as DOM and its utilization would have required measurements of the
13C-labelled DOC pool in the water samples and pore waters and
could not be carried out as part of this study.

Similar to other cold deep-sea sediments (e.g. Woulds et al., 2009), the
fresh phytodetritus was mainly respired, i.e. traced back in DIC in the
overlying water or pore water pools, whereas the assimilation into biomass
was less important. In accordance with Soltwedel et al. (2000), bacteria
clearly displayed the highest biomass in our study area (95 % of the
assessed biotic organic carbon pool), whereas infauna biomass never exceeded
5 % of the total biomass. Therefore, the respiration of the added algae
is, most likely, primarily attributed to bacteria. This corroborates previous
experimental studies (Moodley et al., 2002; Witte et al., 2003b) and is in
agreement with earlier in situ measurements (Donis et al., 2016) and food
web modelling results that assigned 93 % of the respiration to bacteria
(van Oevelen et al., 2011). An increase in bacterial activity was clearly
observed after the deposition of fresh phytodetritus, both in terms of
respiration and enzymatic activity, which is in accordance with earlier
studies (Boetius and Lochte, 1994; Hoffmann et al., 2017). Also, bacterial
fatty acids indicate an uptake of algal-derived carbon, while no significant
increase in abundances was observed even after 14 days (Fig. S4). This may be
related to the relatively short experimental time frame and slow turnover
time of microbial communities in Arctic deep-sea environments – of the order
of 4–5 weeks (Boetius and Lochte, 1994; Hoffmann et al., 2017).

Overall, bacterial community composition was similar to previous reports from
global and Arctic deep-sea sediments (Bienhold et al., 2016; Hoffmann et al.,
2017). The changes in community composition in response to the addition of
phytodetritus were partly consistent with previous studies from the same area
(Hoffmann et al., 2017), e.g. the relative increase in Colwelliaceae
(Gammaproteobacteria) in the Thalassiosira 14-day
experiment. In contrast, there seemed to be little increase in
Bacteroidetes and Flavobacteria, which are usually typical
degraders of complex algal organic material (Hoffmann et al., 2017; Teeling
et al., 2012). However, in the current experiment, a clear response (i.e.
change in community composition) was only observed for the
Thalassiosira 14-day treatment. This may be a consequence of the
less efficient use of Emiliania OM and the lower contribution of
bacteria in the assimilation of carbon in these treatments, especially
considering the relatively short experimental time frame and the
above-mentioned slow doubling times for bacterial communities in Arctic
deep-sea sediments.

It seems that Emiliania OM was initially (4 days and start of
14-day experiment) more respired than Thalassiosira (in 4-day
experiment: 4 % of the added Emiliania OM, of which 3.6 % by
DIC release, as opposed to 2 % of the added Thalassiosira OM),
but this could also be ascribed to dissolution of the inorganic
coccoliths. There was no NH4+ or NOx-
release observed as should co-occur with OM mineralization. This agrees
with a significant contribution of coccolithophorid dissolution to the
observed DIC release. As the lysocline in the Arctic Ocean lies at ∼4000 m water depth (Jutterström and Anderson, 2005), it appears
unlikely for the calcite from the coccoliths to quickly dissolve at our study
site at 2500 m water depth. Nevertheless, Godoi et al. (2009) showed that
the release of CO2 during bacterial respiration can cause the
decrease in the saturation state of seawater in the cell's microenvironment
and may hence favour CaCO3 dissolution.

4.2 Nitrogen cycling

Mineralization of the phytodetritus was also observed in terms of nitrogen.
In deep-sea sediments with low OM content and deep oxygen penetration,
nitrate is expected to be the dominating form of nitrogen recycled
(Brunnegård et al., 2004). Measurements of nitrogen cycling in
oligotrophic deep-sea environments are very scarce (Berelson et al., 1990;
Brunnegård et al., 2004) and to our knowledge do not exist for Arctic
deep-sea sediments. Most of the OM is aerobically mineralized in the upper
centimetre of the sediment in our study area (Donis et al., 2016). Therefore,
the probability that obligate anaerobic processes like denitrification take
place is very low. Hence we assume that denitrification of nitrate does not
occur in the oxidized sediment layer of our experiments and that the observed
accumulation of nitrate in the overlying water is caused solely by
nitrification. In this case, the nitrification rates derived from turnover of
the algal 15N equal 0.35 and 0.38 µmol
15N m−2 d−1 (Emiliania and
Thalassiosira 14-day experiments respectively) and
0.56 µmol 15N m−2 d−1 (Thalassiosira
4-day experiment). To express these rates in terms of algal N nitrified, the
labelling percentage of the total NH4+ concentrations has to be
taken into account. Because 15N-NH4+ was produced
during the release from the algal detritus, the labelling fraction
(15N-NH4+ : total NH4+) in the pore
water increases exponentially (Song et al., 2016). This can be observed in
the upper cm of the sediment where the labelling fraction increased over
time (compare Fig. 1d with Fig. S3; calculated labelling fraction in
Table S2). Taking these labelling fractions of
15N-NH4+ into account, the according nitrification
rates are then 0.74 µmol N m−2 d−1
(Thalassiosira 14-day experiment), 2.1 N m−2 d−1
(Emiliania 14-day experiment) and
4.0 µmol N m−2 d−1 (Thalassiosira 4-day
experiment), an order of magnitude lower than the nitrification rates
observed in other oligotrophic deep-sea areas (Berelson et al., 1990;
Brunnegård et al., 2004). Altogether, only 0.4 %–1.5 % of the
total nitrification would then be attributable to nitrification of the
ammonium released by the algal detritus, which corresponds to the original
addition of algal nitrogen of 0.3 %–0.4 % to the sediment ON pool.
However, these nitrification estimates are based on an addition of fresh
phytodetritus that was heavily (99 %) diluted into the
(labile and refractory) OM pool of the sediment; hence it was an underestimation.

4.3 Assimilation into biomass

Altogether, 0.6 %–6.3 % of the added carbon was assimilated into
biomass. Within this fraction, bacteria seemed to be key players in cycling
freshly added Thalassiosira phytodetritus (∼90 % of the
biological assimilation occurred through bacteria, ∼10 % through
infauna). The share of bacteria was smaller in the Emiliania
experiments (48 %–68 % of the biotic assimilation is accounted for
by bacteria, 32 %–52 % by infauna). This difference is not driven by
a difference in biomass, since the four experiments displayed a similar
bacterial vs. infauna biomass ratio. The absolute assimilation by infauna was
similar in all experiments, whereas the absolute assimilation by bacteria was
∼7 times higher in the Thalassiosira compared to the
Emiliania experiments. This seems to indicate that diatom biomass
was more easily accessible to bacteria compared to coccolithophorids. This
is in contrast with water column studies of Iversen and Ploug (2010), who
found that the degradability of phytoplankton in surface waters depended
partly on the structure of the external mineral protection. In the silica
diatom frustrule, Si–C or Si–O–C interactions are thought to protect
silica from dissolution until the organic matrix is removed by bacteria
(Moriceau et al., 2009 and references therein). Without this organic
protection layer, the diatom frustrule rapidly dissolves in undersaturated
seawater (Ragueneau et al., 2006) ([SiO2] at our study site ∼10µM vs. SiO2 solubility ∼1000µM at
2500 m water depth and 1.4 ∘C, Sarmiento and Gruber, 2013).
Similarly, the calcite matrix of the coccoliths can act as a physical barrier
against bacterial degradation in laboratory experiments (Engel et al., 2009).
However, comparable carbon-specific respiration rates were measured for
aggregates of Emiliania and Skeletonema diatoms, suggesting
similar degradability in laboratory experiments using surface waters (Iversen
and Ploug, 2010). Increased hydrostatic pressure such as in the deep sea
leads to faster dissolution of the coccoliths than in surface waters but in
the presence of natural prokaryotic communities also induces more aggregation
(Riou et al., 2018), which can again offer organic matter protection from
solubilisation and remineralization (Engel et al., 2009). However, silicate
dissolution is reduced when diatoms embedded in sinking aggregates fall
through the water column (Tamburini et al., 2006). It is therefore unclear
whether diatoms should be easier to degrade by bacteria than coccolithophores
when they reach the deep-sea floor. Our data suggest that, in Arctic deep-sea
sediments, Thalassiosira might be more easily degraded by bacteria
than Emiliania. Nevertheless, the state in which the two species
were added at the start of our experiments might differ from the aggregates
and faecal pellets formed during the descent through the water column.

Also noteworthy is the fact that larger organisms dominated the competition
for fresh food: bacterial carbon assimilation never exceeded 1 % of
bacterial biomass, while infauna carbon assimilation reached on average
10 % of their biomass and in Foraminifera even 40 % of their carbon
biomass (Fig. 5). This agrees with other studies showing that bacteria
dominate fresh OM assimilation in sediments only if they are almost devoid of
fauna (Moodley et al., 2005; van Oevelen et al., 2011). However, in systems
where fauna is present, the latter rapidly consume the fresh phytodetritus
(Blair et al., 1996; Levin et al., 1997, 1999; Moodley et al., 2002, 2005;
Witte et al., 2003a, b). This biased ratio indicates that bacteria may be
initially outcompeted by infauna or that carbon becomes available to bacteria
only after it has passed through the guts of fauna (Witte et al., 2003a, b).
Alternatively, sediment reworking (bioturbation) by infauna also
redistributes fresh organic matter deposited at the surface to the deeper
sediment layers, where subsurface bacteria can also access it. Particle
mixing in Arctic sediments is usually limited to the upper 3 cm of the
sediment (Clough et al., 1997; Krauss, 2016; Morata et al., 2015), which fits
with our observations on the increase in subsurface algal-derived OM after
14-day (Fig. 1a) and higher bacterial assimilation in the sediment
subsurface (Fig. 3). However, even after 14 days, Foraminifera still had a
carbon-specific assimilation that was 2 orders of magnitude higher than bacteria,
implying that larger organisms continued to dominate the competition for
fresh OM. This confirms earlier studies showing that foraminifera can be key
players in the early diagenesis of fresh OM at the deep-sea floor (Moodley et
al., 2000, 2002; Nomaki et al., 2005; Woulds et al., 2007). However, these
studies also included the meiofauna fraction of foraminifera
(63–250 µm). Although macrofaunal (> 250 µm)
foraminifera can have a retarded response to phytodetritus inputs compared
to smaller (> 63 µm) foraminifera (Sweetman et al.,
2009), the carbon assimilation rate by macrofaunal foraminifera in this study
is similar to that of smaller foraminifera at Station M (Enge et al., 2011)
and the central basin of Sagami Bay (1449 m) (Nomaki et al., 2005).
Nevertheless, as smaller foraminifera were not analysed here, it may be that
the overall assimilation of this group was still underestimated. Foraminifera
also seemed to prefer Emiliania over Thalassiosira, as seen
in the carbon and nitrogen-specific assimilation. This agrees with the higher
survival rate of planktonic foraminifera in feeding experiments with
Emiliania than with diatoms (Anderson et al., 1979), which was later
related to the higher nutritional value of Emiliania (Pond and
Harris, 1996), but contrasts with observations of abyssal foraminifera at
Station M that showed a preference for nitrogen from diatoms over nitrogen
from coccolithophores (Jeffreys et al., 2013). However, deep-sea foraminifera
demonstrate a variety of dietary preferences (Gooday et al., 2008), and the
species found in this study differ from the ones found in Jeffreys et
al. (2013).

4.4 C : N preference

Measuring the degradation pathways of phytodetritus that is both
13C and 15N labelled allows the preferential assimilation
to be compared in those pools where both isotopes could be traced, i.e.
processed pools (respiration in overlying water, pore water), assimilation by
infauna (assimilation of N into bacterial biomass could not be addressed
based on the measurements performed in this study), and “leftovers”
(sediment OM). The C : N ratio of the added phytodetritus was similar for
both types of algal detritus, with Thalassiosira (13.3) being a
little poorer in nitrogen compared to Emiliania (12.6). The
molar C : N ratio in all processed pools of the short-term experiments was
highest in the Emiliania 4-day experiment (on average 14.3; Table 2)
and considerably lower in the Thalassiosira 4-day experiment (on
average 3.7). This suggests that, in the first stages after the addition of
Thalassiosira, nitrogen is preferably used. This seems to be in
agreement with the nature of the OM leftovers in the sediment, which were
higher in carbon content than the processed pools (C : N ratio 8.7). In
contrast, the preferred use of nitrogen in the Emiliania 4-day
experiment might be masked by the dissolution of the carbonates from the
coccoliths, leading to a higher C : N ratio in overlying and pore water
compared to the C : N in the biomass. The remaining OM therefore contains
less carbon; hence it has a lower C : N ratio compared to the processed
pools. After 14 days, the C : N ratios of both processed and unprocessed
pools in both experiments became increasingly enriched in carbon
(higher C : N ratio). This points again to carbon being increasingly
processed only after the nitrogen content of the available phytodetritus had
strongly been consumed. An exception to these patterns is the C : N ratio
of the infauna in the Emiliania experiments that decreased over
time. Infauna feeding on Emiliania also ingests the coccoliths. As
also observed in the absolute carbon and nitrogen assimilation, it seems that
it takes longer to release the OM from the coccoliths into the guts of
infauna; hence the nitrogen was only accessible later.

We missed certain pools: (a) we could not sample the sediment subsurface
layers of the 4-day Emiliania experiment as deeper layers were lost
upon retrieval of samples from the chambers on board and as such missed the
subsurface processing pathway. However, in the other experiments, this part
accounts for < 10 % of the sum of the total processed carbon and
uncharacterized OM. (b) Unmeasured parts of the carbon and nitrogen budgets
are the DOC and DON pools in the overlying water and pore water. A modelling
study in the same area estimated that > 25 % of the total
carbon input to the food web quickly dissolves into DOC that is taken up and
then respired by prokaryotes (van Oevelen et al., 2011). Based on this
finding, one would expect that the released DOM in the sediment – at some
point but not necessarily in the time frame of the experiments – reappears
in the bacterial biomass or respired pools. (c) We did not consider
meiobenthos < 250 µm, since nematodes, the most abundant
metazoan component of deep-sea meiobenthos, are usually responsible for only
< 1 % of the total mineralization (Ingels et al., 2010).
However, the meiofauna fraction of the foraminifera could have contributed to
the mineralization (Moodley et al., 2002; Nomaki et al., 2005). (d) Bacterial
assimilation of phytodetrital N was not quantified. Assuming a bacterial
C : N ratio of 5 (Goldman and Dennett, 2000) and taking into account that
growth of Arctic deep sea bacteria is N-limited (Boetius and Lochte, 1996),
it can be expected that bacterial N assimilation was up to 5 times lower than
carbon assimilation. This would have doubled the processed share of
Emiliania detrital N after 14 days (from 6 % to 12 % of the
added N) and almost tripled the processed share of Thalassiosira
detrital N after 14 days (from 16 % to 42 % of the added N).
(e) Archaea were not considered, but the experimental duration would probably
have been too short to show a considerable Archaea contribution, as shown for
Thaumarchaeota in shallow Icelandic shelf sediments (Lengger et al., 2014).
Although sequence data suggest that Archaea contribute only 2 %–5 %
to the active members of the benthic prokaryotic community at the study site
(Rapp, 2018), deep-sea Archaea seem to be involved in protein degradation and
carbohydrate metabolism (Li et al., 2015) and deep-sea Archaea from high
latitudes have been shown to be especially sensitive to changes in food
supply (Danovaro et al., 2016).

We added less than we assumed: a substantial part (8 %–60 %) of the
carbon budget concerns uncharacterized particulate OM in the sediment. The
share of this OM increased from 4 to 14 days, which makes the 14-day budgets
more closed. This increase cannot be attributed to a difference in the amount
of algal matter initially added. It may indicate that settlement of the
algal OM to the sediment surface has taken several days and that not the full
amount has been available to the sediment community right from the start.
However, we did not observe increased amounts of POM in the unfiltered
samples of the overlying water. Nevertheless, it cannot be ruled out that
parts of the added OM disintegrated into colloidal particles or were released
from the cells as DOM that took longer to arrive at the sediment.

Finally, the missing part of the budgets and the excess in the
14-day Thalassiosira nitrogen budget might be a result of accumulated errors in integration
procedures due to spatial variability or uncertainties in conversion
factors (Middelburg et al., 2000). For these reasons, the
nature of the missing carbon and nitrogen cannot be fully resolved.

4.6 Long-term implications

Despite its constraints, the presented experiments describe a snapshot of
the potential effects of a change in food quality for Arctic deep-sea
benthos. What could be the long-term implications of a shift in the quality
of food arriving at the deep-sea floor? Our observations suggest that the
degradation of Emiliania-dominated phytodetritus could be less efficient than that of
Thalassiosira-dominated detritus. If this is the case, this shift would affect the
recycling of fresh phytodetritus and the regeneration of nutrients.

Also, the food web structure could be altered: with a shift in food quality,
the share of infauna in OM degradation could increase compared to bacteria.
An increased POC flux to the Arctic deep-sea floor because of reduced sea-ice
cover could in parallel trigger a switch in dominance of benthic organic
matter processing by bacteria to dominance by metazoans, with implications
for the upper food-web levels (Sweetman et al., 2017). Arctic deep-sea
communities could be flexible in their response to new food sources that
accompany climate change and could likely also be influenced by changes in
the amount rather than the type of OM reaching the bottom (Sun et al.,
2007). At our study site, however, there are signs from long-term studies
that both food quality and quantity affect the densities and trophic
diversity of benthic communities (Soltwedel et al., 2016). This calls for
additional in situ experiments in the Arctic deep sea supplying the benthos
with different quantities of OM. Also, OM quality should receive further
attention, since shifts in phytoplankton community structure in Fram Strait
have now turned from diatom-coccolithophore-dominated blooms to
diatom–Phaeocystis pouchetti-dominated blooms (Nöthig et al.,
2015; Soltwedel et al., 2016). Phaeocystis blooms have been shown to
be low in food quality and may impede development in some grazing species
(Breteler and Koski, 2003; Tang et al., 2001). Future investigations on the
effects of altered phytodetritus input to Arctic deep-sea benthos should
therefore also involve the fate of Phaeocystis pouchetti blooms and
their role for benthopelagic coupling in Fram Strait.

FW and FJ designed the experiment; FJ, FW and CB performed the
experiment; UB, CB, HM, ME and CB analysed the samples; UB, GL,
CB and HM analysed the data; and UB wrote the manuscript with contributions from all authors.

We thank the captain and the crew of RV Maria S. Merian expedition
MSM29 for their help during the expedition to the LTER observatory
HAUSGARTEN. We further thank Anja Pappert for culturing the algae, the
MPI-SeaTech technicians for preparing and operating the lander, Christiane
Hasemann for analyses of FDA hydrolysis, Martina Alisch, Rafael Stiens and
Gabriele Klockgether for laboratory analysis of sediment properties,
Jenny Wendt for help with PLFA extractions, Clara Martínez Pérez for
infauna EA–IRMS analyses and Thorsten Dittmar for DOM analyses. Katja
Guilini and Nicolas Van Oostende are acknowledged for fruitful discussions
and Ann Vanreusel for critical reading of the manuscript. We are very
grateful for the constructive feedback on the discussion version of this
manuscript that was provided by Jack Middelburg and an anonymous reviewer.
This work contributes to the framework of the HGF Infrastructure Program FRAM
of the Alfred-Wegener-Institute Helmholtz Center for Polar and Marine
Research. Funding was received from the European Research Council Advanced
Investigator grant 294757, from the Research Foundation – Flanders (FWO
Belgium) to Ulrike Braeckman (grant no. 1201716N).

The article processing charges for this open-access
publication were covered by the Max Planck Society.

Global warming has altered Arctic phytoplankton communities, with unknown effects on deep-sea communities that depend strongly on food produced at the surface. We compared the responses of Arctic deep-sea benthos to input of phytodetritus from diatoms and coccolithophorids. Coccolithophorid carbon was 5× less recycled than diatom carbon. The utilization of the coccolithophorid carbon may be less efficient, so a shift from diatom to coccolithophorid blooms could entail a delay in carbon cycling.